The
figures and, in part, the information
in this section are from “Using
Animals in Intramural Research:
Guidelines for Investigators,”
NIH Animal Research Committee, with
permission of James F. Harwell,
VMD, Chief, Veterinary Resources
Branch, NIH.
Mice
Mice
are usually caught and lifted by
the tail. The tail should be grasped
between its midpoint and the mouse’s
body. With this simple method of
holding, they may be transferred
to another cage or a balance, identified,
examined casually or sex may be
determined. However, such restraint
is not sufficient for treatment
and close examination. For more
effective control, the mouse may
be held by the tail and placed on
a table or other surface, (preferably
one such as a wire cage lid that
the mouse can grasp) and the loose
skin over neck and shoulders grasped
with thumb and fingers. It is necessary
to perform this maneuver expeditiously,
or the mouse may turn and bite.
once the mouse is grasped correctly
the head is adequately controlled.
Restraint is improved if the tail
or the tail and rear legs are held
by the third and little fingers
of the same hand or with the other
hand.
Rats
Rats
are normally lifted by grasping
the whole body with the palm over
back and side with forefinger behind
the head and the thumb and second
finger under opposite axilla. This
extends the rat’s forelimbs
so that they may be controlled (figure
2). Holding with one hand is usually
adequate for control, but the tail,
rear legs or lower part of body
may be held by the other hand for
close control, or treatment or examination.
Young rats may be handled like mice,
when body size does not permit ease
of handling within the hand. Rats
should not be lifted by the tail
as they may spin and strip the skin
from the tail. Rats will bite, and
certain strains are more aggressive
than others (e.g., F344 rats tend
to be more aggressive than Sprague-Dawley),
so care and experience are essential
to rapid handling. Various restraining
devices are available for use with
rats.
Hamsters
Hamsters
that are not used to being handled
may bite, especially when startled. Consequently, they should be approached
gently and with caution until they
become accustomed to being handled
and familiar with the handler. Several
methods may be useful in handling
the hamster. Both hands may be cupped
under the animal to hold the hamster
in the palms. Once a hamster is
lifted in this way, the thumb can
be placed under the chin and the
forefinger around the neck to afford
good control with one hand. Grasping
the loose skin over the neck and
shoulders also provides an effective
method of control with one hand,
however, this skin is very loose
and practice is necessary (Figure
3). It is sometimes easier for the
occasional handler to herd a hamster
into a cup when transferring hamsters
from point to point if detailed
manipulations are not necessary.
Gerbils
Gerbils
are effectively handled by the general
methods indicated for other small
rodents. For rapid handling (as
in transferring from cage to cage),
lifting by the tail near the body
is acceptable. Avoid holding gerbils
near the end of the tail since the
skin near the tip of the tail is
fragile and may slip off. When restraint
of longer duration is required,
gerbils are held much like rats.
Guinea
Pigs
Guinea
pigs seldom bite, but are timid
or easily frightened and usually
make determined efforts to escape
when held. They are best held by
placing the thumb and forefinger
around the neck, with the palm over
the back and the other fingers grasping
the body. When lifting guinea pigs,
one hand should be used to support
the lower part of the body (Figure
4). Special care should be exercised
in supporting the lower part of
the body of pregnant females since
they may become very heavy and awkward
in late pregnancy.
Rabbits
Rabbits
seldom bite, but can inflict painful
scratch wounds, especially with
the hind feet. They should be held
in a way that directs their hind
feet away from the handler’s
body. Grasping the loose skin over
the neck and shoulder with the head
directed away from the holder is
the best method of initial restraint.
When lifting a rabbit, the lower
part of the body must be supported
by the other hand to prevent serious
injury to the rabbit’s back
(Figure 5). Rabbits should never
be restrained or lifted by the ears.
If the rabbit begins to struggle
violently, it should immediately
be placed on a solid surface and
calmed. Struggling frequently leads
to fracture of lumbar vertebrae
and injury to the spinal cord that
may necessitate euthanasia.
During
restraint, rabbits may exhibit sudden
violent efforts to escape and, in
the process, dislodge intravenous
needles, gavage tubes, etc., causing
spills or otherwise endangering
themselves or personnel. Therefore,
complete restraint should be accomplished
before attempting such procedures
and rabbits should never be left
unattended in restraint devices.
Practice in using these devices
can be arranged for through the
OAR.
Other
Species
Recommendations
for handling other species can be
found in a variety references. The
LJFAW Handbook on the Care and Management
of Laboratory Animals (6th Edition,
1987, Poole, T. (ed.), Longman Scientific
and Technical, Avon, England) has
recommendations for handling reptiles,
amphibians, fish, dogs, cats, non-human
primates, ferrets, swine, sheep,
goats, and most other species commonly
used in laboratories.
Administration
of Fluids and Drugs
When
drugs, vaccines, injectable anesthetics
or other agents are to be administered,
one of several different routes
may be selected. The route is governed
by the nature of the agent being
administered, the animal, and the
purpose of the administration, among
other factors.
The
more common routes of administration
used for laboratory animals are
classed as follows:
Substances
may be administered orally by addition
to the food or the drinking water,
by use of a capsule or pill, by
instillation into the mouth using
a mechanical device, such as syringe
or forceps, or by hand. Capsules
or coated pills are rarely used
in rabbits or rodents. When used
in larger animals, capsules or pills
are given by a “pilling gun”
or “balling gun,” or
are placed in the mouth near the
back of the tongue, and the animal
is induced to swallow by stroking
the throat.
Stomach
tubes or gastric feeding needles
are inserted through the mouth into
the stomach or lower esophagus (Figure
7). Care must be taken that the
tube or needle does not enter the
trachea or puncture the esophagus
or stomach. In most cases, introduction
of the tube toward the rear of the
mouth will induce swallowing, and
the tube will readily enter the
esophagus. A violent reaction (coughing,
gasping) usually follows accidental
introduction of the tube into the
larynx or trachea. Flexible or plastic
tubes may be bitten or chewed, and
some care must be taken to prevent
this. With rabbits and some larger
animals, a wooden or plastic dowel
with a hole in the center is held
crossways behind the incisors, and
the mouth held shut by an assistant.
This prevents chewing and permits
entrance of the stomach tube through
the hole in the dowel and on into
the stomach. Rabbits should be placed
in a restraining device before attempting
this procedure, to avoid unnecessary
struggling and injury. A small curved
metal tube with a ball on the end
(feeding needle) is often used with
rodents. Entry normally may be obtained
without anesthesia using hand restraint.
The ball at the end of the tube
prevents trauma to the esophagus
and oral cavity. With the stomach
tube fitted to a syringe or aspirator,
materials may be administered or
withdrawn as required.
Rectal
administration is infrequently used,
however, certain agents are administered
in this manner. Flexible tubing
or inflexible blunt probes or tubes
may be used, but in both cases complete
restraint and gentle procedures
are required. The catheter or probe
must be lubricated with surgical
jelly or petroleum jelly, and entry
should be accomplished slowly to
permit the anus to dilate. Forceable
entry should not be attempted and,
if blockage is encountered, its
cause must be carefully determined
before proceeding.
Parenteral
Parenteral
routes of administration involve
injections into various compartments
of the body. Sites used for collection
of blood from veins may also be
used for intravenous administration.
Intraperitoneal administration is
one of the most frequently-used
parenteral routes in rodents. Other
common locations are the musculature
and subcutis. Materials given intramuscularly
must be in small volumes. Absorption
by this route is more rapid than
from subcutaneous administration.
Regardless of the route used, it
is essential that the subject be
securely restrained to prevent unnecessary
struggling by the animal and to
avoid injury to personnel by dislodged
needles.
The
investigator should know the physiological
and chemical properties of the substance
that he/she plans to inject. Considerable
tissue damage and discomfort can
be caused by irritating vehicles,
drugs or solutions that are cold
when injected into animals. The
use of the foot pad as an injection
site for antigens with or without
adjuvant is discouraged since it
is a needless and painful procedure.
More suitable sites for antigen
injections are subcutaneously in
the axilla or lateral thoracic wall,
deep in large muscle masses, or
into the popliteal lymph node.
The
following outline provides basic
information on equipment and techniques
for parenteral injections in rodents
and rabbits. Demonstration/instruction
sessions can be arranged with the
OAR.
Mouse
Intravenous: Equipment:
27-30 g needle, 1 ml tuberculin
syringe, mouse holder, warming lamp.
The
lateral veins of the tail are the
most frequently-used veins. Best
results are obtained if the tail
is immersed in warm water or the
mouse warmed in the cage with a
warming lamp. The veins can be seen
when the tip of the tail is lifted
and rotated slightly in either direction.
The tip of the needle can be followed
visually as it penetrates the vein.
Trial injection verifies proper
needle placement. Practice is essential.
lntraperitoneal:
Equipment: Syringe and 23 to 27
g, 1/2 to 1-inch needle, preferably
with a short bevel.
The
mouse is grasped as previously described
(Figure 1), and held in dorsal recumbency
in a head-down position. The injection
is made in the lateral aspect of
the lower left quadrant (Figure
8). The use of a short bevel needle
inserted through the skin and musculature
and immediately lifted against the
abdominal wall, aids in avoiding
puncture of the abdominal viscera.
Immobilizing the left leg is also
essential in reducing this risk.
Rapid injection, especially with
a large syringe, may cause discomfort
and tissue damage, and should be
avoided. The maximum volume injected
lP into a 20 gm mouse should not
exceed 2 ml.
Intramuscular:
Equipment: Syringe and 26 to 30
g, 1/2 inch needle.
This
route is usually not used because
of the small muscle mass available
and the danger of damaging vital
structures. However, when it is
used, the back and hind leg muscles
are the usual sites selected. The
maximum volume injected intramuscular
in the mouse should not exceed 0.2
ml.
Subcutaneous:
Equipment: Syringe and 25 to 27
g, 1/2 to 3/4 inch needle. This route is frequently used as
an alternative to intramuscular
injections in the mouse. The site
usually chosen is the area between
the shoulder blades.
Alternatively,
the ventral abdomen is commonly
used; employing restraint demonstrated
in Figure 1. The maximum volume
injected subcutaneously in the mouse
should not exceed 1 .0 ml.
Rats
Intravenous:
Equipment: Depending upon the size
of the rat, needles as large as
20 g and 1 or 1-1/2 in. may be used.
A rat holder and warming lamp are
usually necessary.
The
techniques described for the mouse
apply. In addition, the saphenous
vein on the lateral aspect of the
hind leg may be used. Restraint
must be adequate for the safety
of both the investigator and the
animal. Confinement within a cylindrical
holder is the usual method for restraint.
Light anesthesia with ketamine-xylazine
or CO2 is helpful for restraint.
Prolonged IV administration/sampling
can be accomplished by jugular vein
catheterization, requiring surgical
implantation.
Intraperitoneal:
Equipment: Syringe and 23 to 25
g, 5/8 to 1 inch-needles are recommended.
The
preferred injection site is the
lower quadrant of the abdomen, as
described for the mouse. Restraint
is best accomplished with a second
person holding the rat in a head-down
stretched-out position, or light
anesthesia is recommended.
Intramuscular:
Equipment: 25 to 26 g, 1/2 to 5/8
inch needle and tuberculin syringe.
The
back muscles and hind leg muscles
are used. Precautions to avoid damage
to vital structures must be taken.
Light anesthesia with ketamine-xylazine
or CO2, or the assistance of a second
person as holder, is recommended.
Subcutaneous:
Equipment: Syringe and 23 g, 1 inch
needle is recommended.
The
usual site is between the shoulder
blades. If subcutaneous injections
are done frequently in the same
animal, rotate sites. Again, be
sure to use adequate restraint.
Caution: rat skin is thick and difficult
to penetrate. Care should be taken
to avoid accidental human injections.
Rabbits
Intravenous: Equipment:
20 to 25 g needle of suitable length
with syringe. A
short bevel needle not more than
1 inch long and a syringe of 5 ml
capacity or less is recommended.
A rabbit holder of metal, plastic,
or wood construction is required.
Place
the rabbit in a holder. (Do not
attempt this procedure using manual
restraint.) The marginal ear vein
is used almost exclusively. The
hair over the vein is clipped or
shaved and the skin cleansed with
alcohol or alcohol-iodine before
making the injection. The vein can
be distended by flicking the margin
with the fingers a few times. Pinching
off the vein near the base of the
ear will also help to distend the
vein. Xylene may be used as a vasodilator
by gently rubbing a small amount
of it over the outer surface of
the ear. Xylene is a powerful skin
irritant and extra care is necessary
to clean the ear thoroughly with
soap and water after its use. A
light dose of Innovar-Vet or Acepromazine
will dilate veins and also facilitates
restraint.
Intraperitoneal:
Equipment: 19 or 20 g, 1 or 1-1/2
inch needle with suitable syringe. A flat board or trough with cleats
on the four corners for securing
restraint cords. Four lengths of
soft 1/4 inch rope or 1/2 inch wide
cotton or nylon tape are needed
to tie the legs.
The
rabbit is stretched out on its back
with his legs tied to the corner
cleats. (An assistant may be very
helpful, and chemical restraint
with ketamine is recommended to
reduce struggling.) The abdomen
is clipped and the skin disinfected.
The board is inclined slightly in
a head down position. Injections
are made in the lateral aspect of
the lower abdominal quadrants. Caution
must be taken to avoid puncturing
a distended urinary bladder, the
bowel, or the liver.
Intramuscular:
Equipment: 22 or 23 g, 1 inch needles.
The most frequently-used sites are
the back muscles lateral to the
vertebrae and caudal to the ribs,
or the lateral thigh muscles (more
desirable). If repeated injections
are to be made, rotate sites. The
hair should be clipped and the skin
disinfected. Adequate restraint
is important.
Subcutaneous:
Equipment: 20 to 23 g, 1 inch needle.
The
area most frequently used is between
the scapulae. Clean the skin with
alcohol, clip or part the hair,
and fold the skin over the needle
rather than thrusting the needle
into the skin.
Guinea
Pig
Intravenous: Equipment:
27 to 30 g 1 inch needle.
Several veins are used, but there
is considerable variation among
individuals in their size and usefulness.
Among the veins used are the lateral
metatarsal, cephalic, and saphenous
veins (in which a 27 ga needle is
used) and the marginal ear vein,
which may be used with a very small
(30 ga) needle if the ear is not
pigmented. Chronic indwelling catheters
may be surgically placed in the
jugular vein.
lntraperitoneaI:
Equipment: 19 to 22 g needles with
appropriate size syringe. Restraining
board if working alone.
Light
anesthesia is recommended. Injections
are made in the lower abdominal
quadrant, off the midline, taking
care to avoid the urinary bladder,
bowel and liver.
Intramuscular:
Equipment 20 to 22 g, 1 inch needles
with a 1 to 5 ml syringe.
Intramuscular
injections are not frequently used,
however, if care is taken not to
injure adjacent structures, injections
can be made into the lower back
muscles and lateral muscles of the
thigh (preferred). An assistant
is usually needed.
Subcutaneous:
Equipment: 20 to 22 g, 3/4 to 1
inch needles with appropriate size
syringe.
The
skin of the guinea pig is thick
and tough, especially over the shoulders.
The use of short, heavy gauge needles
are recommended, e.g., 20 g, 1 inch.
Good restraint is necessary to avoid
injury.
Blood
Collection
The
amount of blood needed and other
factors will govern the method and
sites of collection. Table V lists
common blood withdrawal sites in
laboratory animals, and precautions
and requirements for these procedures.
Site
Preparation:
Certain general procedures and precautions
are applicable to methods of blood
collection as well as to administration
of fluids, including injectable
anesthetics. When venipuncture is
required, hair should be shaved
or at least clipped from the site
for better visibility. The area
of injection or incision should
be cleaned with alcohol. Some procedures
will require sedation or anesthesia;
others may be carried out without
anesthesia, provided suitable restraint
is used. In order to better visualize
veins, one of several methods of
dilation may be used: the vessel
may be occluded, the area around
the vessel may be warmed, or xylene
may be swabbed on the site. When
using the rabbit ear or mouse or
rat tail, a low-wattage light bulb
can be used to supply heat. This
also aids by providing additional
light. Because xylene is irritating,
skin swabbed with xylene must be
thoroughly cleaned with alcohol
before drawing blood. Frequently,
brisk rubbing of the skin with a
gauze sponge moistened with alcohol
will produce adequate dilation of
the vessel. Finally, use of certain
sedative agents (eg. Acepromazine
or Innovar-Vet) may produce optimal
vasodilation without use of topical
irritants.
Equipment
Needed:
Needles
of appropriate gauge and length
must be selected with care, especially
for venipuncture. For the tail vein
of mice and rats, the smaller needles
(25 to 30 gauge) should be used.
For other vessels and in other animals,
the suitable size will depend on
size of an animal and site selected
for sampling.
Technique:
Venipuncture - Proper insertion of the needle
into a vein or other part of the
vascular system is normally the
most tedious part of the procedure.
Certain guidelines can be given,
but only practice provides proficiency.
Veins may be expected to roll, collapse,
or shift, making entrance difficult.
A precise, careful introduction
of the needle is best and several
attempts may be required. Starting
at distal sites will allow repeat
attempts more proximally. The needle
is inserted parallel to the vein
and the tip directed into the lumen
along the longitudinal axis. When
withdrawing blood from a vein, aspiration
should be slow so the vessel does
not collapse.
The
marginal ear vein of the rabbit
is useful for collection of small
volumes of blood, and may be used
for intravenous injection. Blood
collection is fairly simple at this
site. The area is shaved and cleaned
with xylene followed by alcohol.
The vein is occluded, the needle
carefully inserted, and blood slowly
withdrawn. Use of a butterfly set
may avoid damage to the vessel if
the animal moves. Gauze held with
pressure over the venipuncture site
for a few minutes will prevent hematomas
from forming. Suitable methods for
collecting larger volumes of blood
from the rabbit central ear artery
include using a 50 ml bottle, or
a plain 20 ga. needle attached to
a silicone-coated tube. 30-40 cc
of blood can be collected in this
manner, but the rabbit must be carefully
restrained and hematomas must be
prevented by direct pressure. For
both methods, prior sedation of
the rabbit will minimize distress
and movement, while promoting vasodilation.
Blood
collection from the jugular vein
is difficult in smaller laboratory
animals such as rodents and rabbits,
but is common in larger animals
such as dogs and cats. In the smaller
species, exposure of the vein under
the skin by surgical cut-down is
usually required if this vein is
to be used.
Withdrawal
of blood and intravenous injection
by way of the tail vein in mice
and rats is possible. The veins
may be seen laterally near the base
of the tail, but good illumination
and dilation will normally be required.
A small blood sample may be collected
by capillary action, using a microhematocrit
tube and small needle placed in
the tail vein.
Blood
may be obtained by cutting off toes
of small rodents or the tip of tails
of mice and rats. It normally amounts
to only a few drops; adequate for
hemoglobin, microhematocrit and
cell counts. Bleeding from the tail
may be increased by warming it in
water at 40o to 5OoC. Animals should
be anesthetized before collections
are made in this way. After cleaning
the tail, 1 to 2 millimeters of
the tip of the tail is cut off and
blood collected. When blood does
not readily appear, there is often
the temptation to “milk”
the tail to induce blood flow. This
must be avoided since it expresses
other body fluids and may yield
an erroneous sample. Clipping toes
or tails are among the least preferable
methods for blood collection.
Cardiac
puncture - Cardiac puncture represents a
practical method of blood collection
from small rodents when more than
a few drops are required. Unfortunately,
this method also carries considerable
risk to the animal, and occasionally
deaths occur. Animals must be anesthetized
and restrained in dorsal recumbency.
The point of strongest heart beat
is determined with the forefinger,
and the needle is then inserted
directly into the heart (Figure
9). Blood should be withdrawn slowly,
and the amount must be limited (up
to about 4 milliliters from adult
rats) unless euthanasia is planned.
Orbital
sinus - Blood collection from the orbital
venous plexus of rats, and orbital
sinus of mice and hamsters is frequently
used. One quarter ml can be repeatedly
collected from mice at weekly intervals
using this technique. Bleeding the
mouse, hamster and rat by the pen-orbital
region requires that the tube be
directed into the orbital sinus
(Figure 10A), which surrounds the
globe. In the mouse, the tube is
inserted into the medial canthus
of the eye and directed caudally
and slightly dorsally. When hamsters
are bled, the capillary tube is
directed caudally to the caudal
venous sinus (Figure lOB). Bleeding
of the rat from the pen-orbital
venous plexus is facilitated when
the capillary tube is directed from
above the globe, and then caudally
toward the major venous anastomosis
between the deep and superficial
veins of the orbit (Figure 10C).
Knowledge of the location of the
venous structures of the orbit of
the mouse, hamster and rat aids
in establishing a successful pen-orbital
bleeding technique. Pressure should
be applied after blood collection
to prevent hematomas. Light anesthesia
is recommended for all pen-orbital
bleeding procedures.
Blood
collection procedures and sites
for dogs, cats, non-human primates,
and livestock species are well described
in several veterinary and technical
texts. For these species, as well
as rodents and rabbits, technical
assistance is available through
OAR.
The
volume of blood removed at any one
time should not exceed 1 .5% of
body weight. Animals should be allowed
at least two weeks between bleedings
that approach this maximum. More
frequent bleeding or collection
of larger volumes necessitates monitoring
hematocrit, hemoglobin and total
plasma protein to detect developing
anemia.
Technical
Assistance Program
Professional
and technical staff of the Office
of Animal Resources are available
to provide assistance or training
to investigators and their technicians
in animal experimental procedures.
The OAR has a collection of training
materials, including audio-visual
training programs, and has information
about people using a wide variety
of experimental procedures at MU.
The OAR can usually point researchers
to a source whose knowledge and
skills are needed. The technical
assistance program provides instruction
in those skills which the investigator
or technician may not be familiar
with or need only sporadically.
Examples include special methods
of euthanasia, conduct of aseptic
surgery, surgical and catheterization
techniques, and methods for specimen
collection, etc. Hands-on workshops
in animal techniques will be periodically
presented as part of this program.
These workshops provide training
in the areas of handling and restraint,
injection and blood collection techniques,
anesthesia, and euthanasia of various
laboratory animals.
The
OAR staff is also available to perform
a variety of animal experimental
procedures on a fee-for-service
basis. This sometimes avoids the
necessity for investigators to hire
and train technicians to do experimental
procedures on animals.
